Silver as a Residual Disinfectant To Prevent Biofilm Formation in Water Distribution Systems
Nadia Silvestry-Rodriguez,1
Kelly R. Bright,2 Donald C. Slack,3 Donald R. Uhlmann,4
and Charles P. Gerba2*
Department of Agricultural and Biosystems Engineering, Room 403, Building 38,
The University of Arizona, Tucson, Arizona 85721,1
Department of Soil, Water and Environmental Science, Room 429, Building 38, The
University of Arizona, Tucson, Arizona 85721,2 Department of Agricultural
and Biosystems Engineering, Room 403, Building 38, The University of
Arizona, Tucson, Arizona 85721,3 Arizona Materials Laboratory,
Department of Materials Science and Engineering, The University of Arizona, 4715 E.
Fort Lowell Road, Tucson, Arizona 857124
Received 1 October 2007/ Accepted 22 December 2007
Biofilms can have deleterious effects on drinking water quality and
may harbor pathogens. Experiments were conducted using 100 µg/liter
silver to prevent biofilm formation in modified Robbins
devices with
polyvinyl chloride and stainless steel surfaces. No
significant
difference was observed on either surface between the
silver
treatment and the control.
The materials used in drinking water distribution systems are readily
colonized by bacteria (5). The
rates of
biofilm formation and release into a distribution system
(DS) can be
affected by many factors (14).
Although
few biofilm organisms pose a threat to humans, many
opportunistic
pathogens are able to survive and proliferate (40).
Chlorination is a commonly used water treatment in the United States and Europe
(41).
Chlorine is also used to provide a residual disinfectant in
the DS
to prevent water recontamination and to maintain the
standards
achieved at the first point of disinfection (4).
Once a
biofilm is established, however, bacteria are more resistant
than
planktonic populations to disinfectants, including chlorine
(16,
20,
32,
44),
and
antibiotics (25).
Factors affecting survival in biofilms in chlorinated water
include
low-nutrient conditions, strain variation, bacterial attachment
to
surfaces with concomitant metabolism changes, and bacterial
encapsulation (1,
19,
43).
Biofilm
growth can lead to pipe corrosion (24,
27),
deterioration in water quality (24)
and
aesthetics (27,
36),
and
other undesirable effects (24).
Chlorine
also produces harmful disinfectant by-products (46),
particularly with high levels of organic matter. Free chlorine
creates problems in older DSs by causing pitting corrosion. Precipitation
of ferric hydroxide accelerates corrosion and represents a
demand on
residual free chlorine aside from that of organic matter (39).
The
identification of safe alternative disinfection methods is
therefore
desirable.
Silver's antimicrobial effect has been demonstrated in numerous
applications
against different types of microorganisms (7,
10).
The
bactericidal efficacy of silver is through its binding to disulfide
or sulfhydryl groups in cell wall proteins (11,
35).
Silver
also binds to DNA (38).
Through
these binding events, metabolic processes are disrupted,
leading to
cell death (21).
Silver has been reported to delay or prevent the formation of
biofilms
in medical catheters (8, 13, 15, 33),
prosthetic heart valves (3, 17),
vascular
grafts, and fracture fixation devices (6, 9).
Silver has
also been used in water filters (31),
cooling
towers (22),
and DSs (23,
26,
29).
Silver
exerts its antimicrobial effect by progressive elution from
the
devices.
Silver is effective against planktonic bacteria (34)
and has
been used for water disinfection in Europe
(18,
31).
In
addition, silver, in combination with copper, has proven
effective
against Legionella pneumophila in hospital DSs for
more than
a decade (37).
Silver is
not believed to react with most organics in DSs or to
produce toxic
by-products (46).
The objective of this study was to determine if silver
inhibits
biofilm formation on two very different surfaces to
evaluate its
potential as a residual disinfectant in DSs.
Tucson municipal tap water
(Table 1)
(groundwater
source) was dechlorinated by passage through a PUR
activated-carbon
filter (Procter & Gamble, Cincinnati , OH).
Two 10-liter tanks were
filled with dechlorinated water containing 0.5 mg/liter
humic acid
(Sigma-Aldrich, St. Louis, MO) as a source of
organic matter since,
unlike surface water, groundwater usually has low organic levels
(2).
The total
organic carbon of water sources ranges from 0.5 to >10
mg/liter (2)
(test waters
averaged 0.43 mg/liter total organic carbon).
TABLE 1. Quality of the water used in the present study
|
Value
|
Chlorine concn (mg/liter)
|
Hardness (mg/liter CaCO3)
|
Sodium concn (mg/liter)
|
Temp (°C)
|
Total dissolved solids (mg/liter)
|
pH
|
|
|
|
Avg
|
0.74
|
140
|
45
|
27.22
|
317
|
7.84
|
|
Lower
|
0.46
|
64.56
|
24.6
|
20.44
|
173.8
|
7.38
|
|
Upper
|
1.15
|
227.84
|
57.3
|
32.5
|
463.9
|
8.17
|
|
In one tank, a final silver concentration of 100 µg/liter was
achieved by adding silver nitrate (Sigma-Aldrich, St. Louis , MO).
This amount is deemed safe for human consumption by the World
Health
Organization (45)
and the Environmental Protection Agency (http://www.epa.gov/safewater/mcl.html).
This concentration was confirmed by using an ELAN DRC-II
(Perkin-Elmer Life Sciences, Shelton , CT).
Experiments were conducted at room temperature (24°C).
Tanks were
placed in line by using silicone tubing with a cassette pump
(Manostat; Barnant, New York, NY) to
supply a constant water
flow (tanks were replenished daily). Water from each tank was
pumped
through two separate modified Robbins devices (LPMR-25; Tyler
Research, Edmonton ,
Canada ).
The first of these had 25
sampling ports outfitted with stainless steel coupons, and
the second
had polyvinyl chloride (PVC) coupons. These surfaces are
common in
DSs and were chosen to ascertain how dissimilarities in
bulk or
surface chemistry, microstructure, and stiffness would
affect
interactions with silver.
Experiments were conducted with a constant water flow (0.41
liter/h).
Three randomly spatially distributed coupons were removed
from each
device at 0, 1, 7, 15, 23, 29, and 36 days. Biofilms were
scraped
from the coupons with a sterile spatula and placed in 1 ml
of D/E
neutralizer (Difco, Sparks, MD) to inactivate
the
silver. Samples were serially diluted in saline (0.85%
NaCl) and
enumerated via spread plating on R2A agar (Difco). Plates
were
incubated at room temperature for 5 days. The number of CFU
of
heterotrophic plate count bacteria per square centimeter was
determined. Analysis of variance was conducted to compare the
treatments to controls by using STATA/SE 9.1 (Stata Corp., College Station, TX).
The results for biofilm formation on PVC and stainless steel
surfaces
are presented in Tables 2 and 3,
respectively.
Despite biofilms forming more rapidly in some cases in
controls,
there was no significant difference (P0.05)
found between the silver treatment
and the control with either test surface. There was also no
significant difference between the two surfaces. Therefore, the
nature of the biofilm, not the surface properties, was responsible
for
silver's lack of effectiveness. This has been observed with other
substances, such as antibiotics, where biofilm growth is
independent
of the underlying biomaterial substrate (28).
In one tank, a final
silver concentration
of 100 µg/liter was achieved by adding silver nitrate
(Sigma-Aldrich, St. Louis ,
MO). This
amount is deemed
safe for human consumption by the World Health Organization
(45)
and the
Environmental Protection Agency (http://www.epa.gov/safewater/mcl.html).
This concentration was confirmed by using an ELAN DRC-II
(Perkin-Elmer Life Sciences, Shelton, CT).
Experiments were conducted at room temperature (24°C).
Tanks were
placed in line by using silicone tubing with a cassette pump
(Manostat; Barnant, New York, NY) to
supply a constant water
flow (tanks were replenished daily). Water from each tank was
pumped
through two separate modified Robbins devices (LPMR-25; Tyler
Research, Edmonton ,
Canada ).
The first of these had 25
sampling ports outfitted with stainless steel coupons, and
the second
had polyvinyl chloride (PVC) coupons. These surfaces are
common in
DSs and were chosen to ascertain how dissimilarities in
bulk or
surface chemistry, microstructure, and stiffness would
affect
interactions with silver.
Experiments were conducted with a constant water flow (0.41
liter/h).
Three randomly spatially distributed coupons were removed
from each
device at 0, 1, 7, 15, 23, 29, and 36 days. Biofilms were
scraped
from the coupons with a sterile spatula and placed in 1 ml
of D/E
neutralizer (Difco, Sparks , MD) to inactivate
the
silver. Samples were serially diluted in saline (0.85%
NaCl) and
enumerated via spread plating on R2A agar (Difco). Plates
were
incubated at room temperature for 5 days. The number of CFU
of
heterotrophic plate count bacteria per square centimeter was
determined. Analysis of variance was conducted to compare the
treatments to controls by using STATA/SE 9.1 (Stata Corp., College Station, TX).
The results for biofilm formation on PVC and stainless steel
surfaces
are presented in Tables 2 and 3,
respectively.
Despite biofilms forming more rapidly in some cases in
controls,
there was no significant difference (P0.05)
found between the silver treatment
and the control with either test surface. There was also no
significant difference between the two surfaces. Therefore, the
nature of the biofilm, not the surface properties, was responsible
for
silver's lack of effectiveness. This has been observed with other
substances, such as antibiotics, where biofilm growth is
independent
of the underlying biomaterial substrate (28).
TABLE 2. Effect of silver on biofilm formation on PVC surfaces
|
Day
|
Mean no. of CFU/cm2
± SDa
|
|
Silver treatment
|
Control
|
|
|
|
1
|
9.4 x 10–3
± 4.8 x 10–3
|
1.1 x 10–2
± 1.0 x 10–2
|
|
|
7
|
9.9 x 101
± 6.8 x 101
|
6.4 x 103
± 9.0 x 103
|
|
|
10
|
8.8 x 101
± 9.2 x 101
|
7.0 x 102
± 8.1 x 102
|
|
|
23
|
5.5 x 101
± 1.7 x 101
|
1.0 x 102
± 7.5 x 101
|
|
|
29
|
8.9 x 103
± 1.3 x 104
|
1.6 x 104
± 2.2 x 104
|
|
|
36
|
7.3 x 103
± 9.9 x 103
|
4.1 x 104
± 5.6 x 104
|
|
|
a The results shown are means
of triplicate
samples.
TABLE 3. Effect of silver on biofilm formation on stainless steel
surfaces
|
Day
|
Mean no. of CFU/cm2
± SDa
|
|
Silver treatment
|
Control
|
|
|
|
1
|
3.7 x 10–2
± 1.7 x 10–2
|
7.7 x 10–2
± 1.5 x 10–2
|
|
|
7
|
1.1 x 102
± 5.6 x 101
|
6.1 x 103
± 8.4 x 103
|
|
|
10
|
1.5 x 102
± 8.1 x 101
|
1.0 x 104
± 1.4 x 104
|
|
|
23
|
2.9 x 102
± 3.2 x 102
|
9.1 x 101
± 5.1 x 101
|
|
|
29
|
2.5 x 104
±3.9 x 104
|
1.7 x 104
± 2.4 x 104
|
|
|
36
|
6.1 x 103
± 8.3 x 103
|
1.5 x 104
± 2.0 x 104
|
|
|
a The results shown are means
of triplicate
samples.
This ineffectiveness of silver on biofilm bacteria stands in marked
contrast to silver's effect on planktonic bacteria in previous
studies (34).
This difference likely reflects the complexing of silver
cations
with the anionic polysaccharide constituent of biofilms.
Biofilms
can sequester minerals and metals from the liquid phase
with which
they are in contact (12).
In
particular, the exopolysaccharides of gram-negative
bacteria play an
important role in metal biosorption. The binding affinity
depends
largely on the cation size/charge ratio, the bacterial
polysaccharide charge, the pH, the physical state of the
biofilm,
etc. (42).
Similar phenomena have been demonstrated for cationic
antibiotics (e.g.,
polymyxin B) that bind to the lipid A portion of lipopolysaccharides
in
gram-negative bacteria (30).
The silver concentrations measured in tank effluents (sampled
prior
to entering Robbins devices) ranged from 90 to 122 µg/liter,
whereas
the system effluents (collected from the distal end of
Robbins
devices) ranged from 14 to 20 µg/liter, indicating that
most of the
silver was likely being absorbed by biofilms. With higher
silver
concentrations or longer exposure times, it should be
possible to
exceed the biofilm absorption capacity; silver would then
inhibit
biofilm development. Calculations are under way to
elucidate the
relationship between biofilm characteristics and the silver
ion
concentration needed to produce net ions for eliminating
the
bacteria in biofilms.
This study was supported in part by the University
of Arizona's National Science Foundation Water Quality Center.
Corresponding author. Mailing address:
Department of
Soil, Water and Environmental Science, The University of Arizona,
Building 38,
Room 429 Tucson, AZ 85721. Phone: (520) 621-6906. Fax: (520) 621-6163.
E-mail: gerba@ag.arizona.edu
Published ahead of
print on 11 January 2008.
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Applied and Environmental
Microbiology, March
2008, p. 1639-1641, Vol. 74, No. 5
0099-2240/08/$08.00+0 doi:10.1128/AEM.02237-07
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